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First published online March 17, 2006
Journal of Experimental Biology 209, 1274-1284 (2006)
Published by The Company of Biologists 2006
doi: 10.1242/jeb.02093
Effects of acclimation temperature and cadmium exposure on cellular energy budgets in the marine mollusk Crassostrea virginica: linking cellular and mitochondrial responses
1 Biology Department, University of North Carolina at Charlotte, 9201
University City Blvd, Charlotte, NC 28223, USA
2 Carolinas Medical Center, Cannon Research Center, 1542 Garden Terrace,
Charlotte, NC 28203, USA
* Author for correspondence (e-mail: insokolo{at}uncc.edu)
Accepted 12 January 2006
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Key words: mitochondrial respiration, cellular respiration, energy budget, mitochondrial membrane potential, proton leak, protein synthesis, cadmium, temperature, bivalve
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; Baird et al.,
1990). The resulting energy deficit can have adverse effects on
survival and performance of organisms and on the long-term persistence of
their populations in the stressful environment. To date, case studies
analyzing stress-induced changes of energy budgets have mostly focused on
single environmental stressors (Widdows,
1978; Dorigan and Harrison,
1987; Koehn and Bayne,
1989; Li et al.,
2002), whereas effects of multiple stressors are not well
understood. Because multiple stresses often have non-additive effects on
physiology, analysis of environmentally relevant combinations of stressors is
important to obtain a realistic picture of the impact of stress on animal
bioenergetics in nature.
Trace metals (such as cadmium) and temperature are common stressors in
estuaries, and their importance is increasing due to global climate change and
accumulation of persistent metal pollutants in coastal habitats
(Helmuth et al., 2002;
GESAMP, 1987). Cadmium (Cd) is
a toxic metal that acts as a potent inhibitor of mitochondrial function in a
variety of plant and animal models at concentrations as low as
10–6 mol l–1
(Kesseler and Brand, 1994;
Kesseler and Brand, 1995;
Korotkov et al., 1999;
Dorta et al., 2003;
Sokolova, 2004 and references
therein). Typically, Cd exposure in vitro results in decreased
mitochondrial efficiency, reduced rate of ATP synthesis and progressive
uncoupling (for a review, see Brierley,
1977; Byczkowski and Sorenson,
1984; Miccadei and Floridi,
1993).
Our recent studies showed that elevated temperatures strongly enhance the
adverse effects of Cd on mitochondrial ATP synthesis and coupling in a model
marine poikilotherm, Crassostrea virginica, suggesting synergism
between these two environmental stressors
(Sokolova, 2004; A. S.
Cherkasov, A. H. Ringwood and I. M. Sokolova, manuscript submitted for
publication). Earlier studies have also shown that elevated temperatures and
exposures to metals may result in elevated standard metabolic rates (SMR) in
poikilotherms (Barber et al.,
1990; Rowe, 1998;
Rowe et al., 1998;
Hopkins et al., 1999;
Rowe et al., 2001;
Willmer et al., 2000;
Lannig et al., 2006). This
suggests that poikilotherms exposed to elevated temperature and toxic metals
may face a dilemma of elevated energy demand combined with reduced aerobic
capacity to produce ATP. However, mechanisms underlying an increase in SMR in
metal-exposed poikilotherms are not well understood and it is not known
whether elevated energy demand can be partially compensated in vivo
(e.g. by increases in mitochondrial abundance or efficiency). A comprehensive
analysis of the combined effects of environmental temperature and Cd exposure
on energy demand and aerobic capacity for energy supply will provide key
information for mechanistic understanding of metabolic effects of these
stressors and will further our knowledge of the role of bioenergetics in
stress tolerance.
Eastern oysters, C. virginica Gmelin (Bivalvia: Ostreidae), are a
useful model for studies of the interactive effects of temperature and Cd
stress on the cellular energy budget of poikilotherms. Oysters are exposed to
varying Cd concentrations in their habitats and have an ability to accumulate
Cd in soft tissues to concentrations exceeding the environmental levels by
orders of magnitude (Roesijadi,
1996). Like all intertidal organisms, oysters may experience
extreme temperature fluctuations in their habitats, with a change in body
temperature as large as 20°C within a few minutes during summer low tides
and up to 35°C during more gradual seasonal variation in ambient
temperatures (Helmuth et al.,
2002; I.M.S., unpublished data). These temperature changes may
strongly affect the SMR of oysters
(Shumway, 1996;
Hutchinson and Hawkins, 1992;
Lannig et al., 2006) as well
as the sensitivity of their mitochondria to Cd
(Sokolova, 2004; A. S.
Cherkasov, A. H. Ringwood and I. M. Sokolova, manuscript submitted for
publication), thereby increasing the potential for interactive effects of
those stressors on both the demand side and the supply side of the energy
budget.
The aim of our study was to examine the effects of acclimation temperature and Cd exposure on cellular energy budget and mitochondrial capacity in C. virginica and to analyze which parts of the cellular energy demand are most strongly affected by temperature and Cd stress. We have studied the effects of temperature (12, 20 and 28°C) and exposure to Cd (50 µg l–1) on the energy budget of oysters using oxygen demand for ATP turnover, protein synthesis, mitochondrial proton leak and non-mitochondrial respiration in isolated cells as demand-side endpoints and mitochondrial oxidation capacity, abundance and fractional volume as supply-side endpoints. This study, for the first time, provides evidence that Cd stress adversely affects both demand and supply sides of the cellular energy balance in C. virginica and that Cd-induced metabolic costs are mostly due to the elevated rates of protein synthesis.
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. The study sites have very low background
concentrations of pollutants (Mallin et
al., 1999; J. Swartzenberg, personal communication). Animals were
transported to the University of North Carolina at Charlotte within 8 h and
placed in tanks with artificial seawater (SW) (Instant Ocean®; Kent
Marine, Acworth, GA, USA) at 15±1°C and 30±0.5.
Oysters were allowed to recover for 3–5 days, and temperature in the
tanks was then gradually changed to reach the required experimental
temperatures (12, 20 and 28°C). The rate of the temperature change was
<2°C day–1, and the duration of the pre-acclimation
period was 8–10 days for all treatments.
After the preliminary acclimation, half of the experimental tanks were
randomly selected, and Cd was added to the nominal concentration of 50 µg
l–1. The remaining tanks were used as controls. Oysters were
incubated for 20 or 40 days at 50 µg l–1 Cd (Cd-exposed
oysters) or in clean artificial seawater (controls) at each temperature. To
avoid pseudoreplication, at least two replicate tanks were set up for each
treatment. Oysters were fed daily with a commercial algal blend (2 ml per
oyster) containing Nannochloropsis, Tetraselmis and
Isochrysis spp. ranging in size from 2 to 15 µm (PhytoPlex©;
Kent Marine). Water was changed every other day. In order to avoid Cd
depletion in Cd-exposed tanks, a static-renewal design was used, with cadmium
supplementation to the nominal concentration of 50 µg l–1
during each water change. Cd levels were measured in water samples at least
twice a week immediately before and 1 h after water changes. There were no
differences in Cd levels before and after water changes, indicating that the
maintenance conditions were adequate to prevent Cd depletion. The average Cd
concentration in experimental tanks was 42.9±7.10 µg
l–1 (mean ± s.d., N=37), and Cd levels in the
control tanks were below the detection limits of the method used (0.5 µg
l–1). It is worth noting that Cd concentrations in our
experimental exposures (50 µg l–1) were at the upper end
of Cd concentrations found in polluted estuaries
(Crompton, 1997;
Hackney et al., 1998).
However, due to relatively short exposure times, the tissue Cd burdens in our
experiments were well within the range found in oysters from polluted sites in
the nature (see Results below and compare with
Roesijadi, 1996 and references
therein). This indicates that our exposure regime resulted in environmentally
realistic tissue Cd burdens and thus that physiological changes observed in
response to Cd exposure in our experiments are environmentally relevant.
Chemicals
Type I collagenase was purchased from Worthington Biochemical Corporation
(Lakewood, NJ, USA), and TMRM (tetramethylrhodamine methyl ester) was from
Molecular Probes (Invitrogen, Carlsbad, CA, USA). All other chemicals were
purchased from Sigma Aldrich (St Louis, MO, USA) or Fisher Scientific
(Suwanee, GA, USA) and were of analytical grade.
Mitochondrial oxidation
Mitochondria were isolated from gills of control and Cd-exposed oysters
after 20 days of exposure at 12, 20 and 28°C. In oysters, gills are a
primary site of uptake of trace metals, which are characterized by early
accumulation of Cd in the mitochondrial fraction
(Sokolova et al., 2005b), and
therefore this tissue is an appropriate model to study effects of Cd on
mitochondria. Isolation was performed as described previously
(Sokolova, 2004).
Mitochondrial oxygen uptake was measured in 1 ml water-jacketed chambers
using Clarke-type oxygen electrodes (Qubit Systems, Kingston, ON, Canada) at
the respective acclimation temperatures (12, 20 and 28°C). Mitochondrial
assay conditions were as described previously
(Sokolova, 2004), and
calibration of oxygen electrodes, data acquisition and rate of oxygen
consumption (
O2)
calculations were performed as described previously
(Sokolova et al., 2005a).
Succinate was used as a substrate at saturating amounts (10–15 mmol
l–1) in the presence of 5 µmol l–1 of
rotenone. Maximal respiration rates (state 3), indicative of the maximum
capacity for ATP synthesis in mitochondria, were achieved by addition of
200–300 nmol ADP, and state 4 respiration was determined in
ADP-conditioned mitochondria as described previously
(Chance and Williams, 1955).
State 4+ respiration was determined as oxygen consumption rate after addition
of 2.5 µg ml–1 of the ATPase inhibitor oligomycin. State
4+ respiration in the presence of oligomycin is considered as a good upper
limit estimate of mitochondrial proton leak measured at high mitochondrial
membrane potential (Brand et al.,
1994). All assays were completed within 2 h of isolation of the
mitochondria. Preliminary experiments have shown that there was no change in
mitochondrial respiration or coupling during this period. Respiration rates
were corrected for electrode drift and non-mitochondrial respiration (see
Sokolova, 2004) and expressed
as natom O min–1 mg–1 mitochondrial protein.
Respiratory control ratio (RCR) was determined as a ratio of state 3 over
state 4 respiration (Estabrook,
1967).
Mitochondrial membrane potential (MMP)
MMP was determined as described in Cherkasov et al. (in review). Briefly,
mitochondrial suspensions were diluted to 2 mg ml–1
mitochondrial protein in the standard assay medium (AM) containing 0.5 µmol
l–1 of the potentiometric dye TMRM, 20 mmol
l–1 succinate and 5 µmol l–1 rotenone.
Preliminary studies have shown that 0.5 µmol l–1 TMRM does
not affect respiration of oyster mitochondria (data not shown). TMRM
fluorescence in mitochondrial suspension was measured under constant stirring
at an excitation wavelength of 573 nm and emission wavelength of 590 nm
(excitation and emission slits 10 nm) using a fluorescence spectrophotometer
(Hitachi Ltd, Tokyo, Japan). The mitochondrial uncoupler CCCP (carbonyl
cyanide-chlorophenyl hydrazone; 50 µmol l–1) was added to
the mitochondrial suspension to collapse the membrane potential, and
fluorescence was measured again. The degree of quenching of TMRM fluorescence
by energized mitochondria at 573 nm excitation wavelength
(1/F573) was used as an index of MMP and was normalized to
the fluorescence level in CCCP-uncoupled mitochondria. Scaduto and Grotoyohann
have previously demonstrated that the degree of quenching of TMRM fluorescence
at 573 nm is directly and linearly proportional to MMP
(Scaduto and Grotoyohann,
1999).
Mitochondrial abundance and fractional volume
Hepatopancreas and gill tissues (1–2 mm3) were fixed in
2.5% glutaraldehyde solution in 0.1 mol l–1 cacodylate buffer
(pH 7.5) containing 150 mmol l–1 NaCl and 300 mmol
l–1 sucrose. Post-fixation in 1% osmium tetroxide buffered
solution was followed by the standard dehydration procedure with increasing
concentrations of ethyl alcohol and acetone
(Hayat, 2000). Tissues were
embedded in Embed-812–Araldite mixture, and an Ultracut ultramicrotome
(Ultracut UCT, Deerfield, IL, USA) was used to cut ultra-thin sections
(60–90 nm). Sections were mounted on 200-mesh copper grids and
double-stained with uranyl acetate and lead citrate according to a standard
protocol (Hayat, 2000).
Sections were analyzed with a transmission electron microscope (Philips CM 10;
Philips Export B.V., Eindhoven, Netherlands) at 60 kV.
Mitochondrial numbers per unit area were calculated using the unbiased
stereology approach on randomly chosen sections of oyster gill and
hepatopancreas cells as follows:
 | (1) |
Q is the sum of
mitochondria counted within all frames and Nf is the
number of frames counted (Howard and Reed,
1998).
The fractional volume of mitochondria in gill and hepatopancreas tissue was
estimated using multi-purpose combined point grid (ratio of fine to coarse
points = 25:1) as described previously
(Howard and Reed, 1998).
Fractional volume [Vv(Y, ref)] was determined as:
 | (2) |
Cell isolation and respiration rates
Cells were isolated from gills and hepatopancreas of control and Cd-exposed
oysters after 20 or 40 days at 12, 20 and 28°C. Oyster shells were scraped
and surface cleaned with 1% bleach. Gills or hepatopancreas tissues from
3–5 oysters were pooled on ice in 5 ml of the digestion buffer
containing 24.72 g l–1 NaCl, 0.68 g l–1 KCl,
1.36 g l–1 CaCl2.2H2O, 0.18 g
l–1 NaHCO3 and 30 mmol l–1 Hepes
at pH 7.5. Tissues were minced and washed twice with 10 ml of the digestion
buffer. Tissue fragments were digested for 10 min at room temperature in
0.125% trypsin in balanced Hank's solution (Fisher Scientific, Suwanee, GA,
USA) adjusted to 720 mOsm with sucrose. Tissue fragments were carefully
triturated to release cells and washed twice with the digestion buffer. The
supernatant was filtered through 100 µm sterile nylon mesh and centrifuged
for 10 min at 400 g to pellet the cells. Cells were washed
three times in cell suspension medium (CSM), containing 24.72 g
l–1 NaCl, 0.68 g l–1 KCl, 1.36 g
l–1 CaCl2.2H2O, 0.18 g
l–1 NaHCO3, 4.66 g l–1
MgCl2.6H2O, 6.29 g l–1
MgSO4.7H2O, 15 mmol l–1 glucose and 30
mmol l–1 Hepes at pH 7.5, and re-suspended in 2 ml of the
same medium. Tissue fragments remaining from the trypsin digestion were
additionally digested with 0.125% of Type I collagenase (Worthington
Biochemical Corporation) dissolved in Mg-free digestion buffer for 40 min at
room temperature, triturated to release cells, washed twice, and filtered
through 100 µm sterile nylon mesh to remove undigested tissue. Cells were
collected by centrifugation as described above, washed three times and
re-suspended in 2 ml of the CSM. Cell suspensions from both digestions were
pooled, enumerated using a hemacytometer, and cell density adjusted to
10x106 cells ml–1. Cell viability was
determined using a standard Trypan Blue exclusion assay and was found to be
>90% (mean 95.9±1.10%).
Cellular respiration was determined in 1 ml water-jacketed chambers using
Clarke-type oxygen electrodes (Qubit Systems, Kingston, ON, Canada) at the
respective acclimation temperatures (12, 20 and 28°C). Total
O2, and
O2 in the presence of 3
µg ml–1 oligomycin (to inhibit mitochondrial
F0, F1-ATPase), 100 µmol l–1
cycloheximide (to inhibit cytosolic protein synthesis) and 100 µmol
l–1 KCN and 200 µmol l–1
salicylhydroxamic acid (SHAM) (to inhibit mitochondrial respiration), were
determined. Cycloheximide is widely used as an inhibitor of cytoplasmic
protein synthesis in ectotherms including marine mollusks
(Giuditta et al., 1968;
Alkon et al., 1987;
Fuery et al., 1998; Joyner and
Peyer, 2003). In our preliminary studies, we tested different cycloheximide
concentrations and found that concentrations between 75 and 750 µmol
l–1 showed a similar and consistent degree of inhibition of
cellular O2 (by
10–15%). This agrees with earlier studies showing that these levels of
cycloheximide specifically and effectively block protein synthesis in isolated
molluscan cells and tissues (Giuditta et
al., 1968; Alkon et al.,
1987). Effects of cycloheximide on cell respiration in oysters
were maximal after 1 h of incubation, with no further decrease in
O2 up to 5 h of incubation
(data not shown). Therefore, in all further experiments we incubated cells
with 100 µmol l–1 of cycloheximide for 1 h on ice. Control
cells were incubated on ice for the same time without cycloheximide addition.
Pilot experiments showed that there was no change in cell viability or
O2 during this incubation.
Cellular responses to oligomycin, KCN and SHAM were immediate (within a few
minutes). All respiration rates were corrected for electrode drift. Oxygen
demand for different cellular processes was calculated as shown in
Table 1.
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Cadmium determination
Gill samples were freeze-dried and digested in Teflon bottles with 52.5%
nitric acid (trace metal grade; Fisher Scientific) using 3–4 cycles of
microwave heating and cooling until the tissues were fully digested. Cell
suspensions were mixed 1:1 with 70% nitric acid and digested as described
above. Water samples were acidified with 0.7% nitric acid, incubated overnight
on a shaker at room temperature and diluted 1:5 with deionized water. Cd
concentrations were determined with an atomic absorption spectrometer (Perkin
Elmer AAnalyst 800; Norwalk, CT, USA), equipped with a graphite furnace and
Zeeman background correction. NIST oyster tissue (1566b; National Institute of
Standards and Technology, Gaithersburg, MD, USA) was analyzed with the samples
to verify the metal analyses; the percent recoveries over all batches were
94.6±6.6% (mean ± s.d.).
Protein concentrations
Protein concentrations in mitochondrial or cell suspensions were measured
using a modified Biuret method with 1% Triton-X added to solubilize the
mitochondria (Bergmeyer, 1985).
Bovine serum albumin (BSA) was used as the standard. Protein content was
measured for each batch of the isolation medium and subtracted from the total
protein content of the mitochondrial or cell suspensions.
Statistics
General linearized model (GLM) analyses of variance (ANOVAs) were used to
test the effects of exposure duration (random effects), and acclimation
temperature and cadmium exposure (fixed effects) after testing the assumptions
of normality of data distribution and homogeneity of variances. Mitochondrial
abundance and fractional volumes were analyzed using repeated-measures ANOVA
with tissue sections nested within individual oysters (random effect) and
tissue, acclimation temperature and cadmium exposure as fixed effects. Dunnett
tests were used for post-hoc comparisons, and LSD (least squared
difference) tests for planned comparisons of sample means as appropriate.
Preliminary tests were run to analyze the effect of replicate tanks within
each treatment on the studied variables (data not shown). No significant
differences were observed between the tanks and the data were pooled to
increase the power of analysis. Statistical analyses were performed using SAS
9.1.3 software (SAS Institute, Cary, NC, USA). Differences were considered
significant if the probability for Type II error was less than 0.05, and
Bonferroni correction was used to adjust significance levels for multiple
ANOVA tests.
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Cellular O2 increased
with increasing temperature (Fig.
1B). In controls,
O2 of gill cells was
notably higher than in hepatopancreas at 12 and 20°C but not at 28°C
(Fig. 1B). Non-mitochondrial
respiration accounted for 12–15% and for 16–31% of total
O2 in gill and
hepatopancreas cells, respectively. In gills, temperature acclimation had no
effect on the proportion of non-mitochondrial oxygen consumption
(F2,70=0.85, P=0.43). In hepatopancreas cells,
the proportion of non-mitochondrial respiration decreased with increasing
temperature, from 31% in 12°C-accimated oysters to 16% in the
28°C-acclimated group (F2,69=15.64,
P<0.001, N=6–12). Cd exposure had no effect on
non-mitochondrial oxygen consumption (P>0.10).
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O2, although this elevation
was only statistically significant in gills and hepatopancreas cells at
12°C and gill cells at 20°C (Fig.
2A–C). Similarly, oxygen demand for ATP turnover
significantly increased in response to Cd exposure at 12 (in gills) and
20°C (in both tissues) but not at 28°C
(Fig. 2D–F). Energy
demand for protein synthesis was strongly enhanced by Cd exposure, especially
at 12 and 20°C (Fig. 3A-C).
In 12°C- and 20°C-acclimated oysters, Cd-induced increase in protein
synthesis was significant in gill and hepatopancreas cells, whereas in
28°C-acclimated ones it was only significant in hepatopancreas. On
average, Cd exposure resulted in a 2–2.5-fold increase in protein
synthesis rates after 40 days of exposure, with the exception of gill cells at
12°C, where protein synthesis rates in Cd-exposed oysters were six times
higher than in the respective controls. Rates of mitochondrial proton leak
were elevated in gill and hepatopancreas cells of oysters after prolonged (40
days) Cd exposure at 12°C (Fig.
3). At 20 and 28°C there were no significant changes in proton
leak rates with Cd exposure (Fig.
3E,F).
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O2 due to
protein synthesis) strongly increased with Cd exposure. Thus, in gill cells
from control oysters, protein synthesis was responsible for 7–23% of
total ATP turnover rate, whereas in oysters exposed to Cd for 40 days this
proportion increased to 41–48%. Similarly, in hepatopancreas cells from
control oysters, protein synthesis accounted for 23–49% of ATP turnover,
whereas in Cd-exposed oysters this proportion rose to 54–71%
(Fig. 4). The proportion of
O2 due to proton leak or
ATP turnover did not change with temperature acclimation or Cd exposure
(Fig. 4). Proton leak accounted
for 21–33% of total
O2 in gills and for
23–37% in hepatopancreas. Oxygen demand for ATP turnover was
53–65% of total O2 in
gill cells at all studied temperatures and in hepatopancreas cells at
28°C. At 12°C and 20°C, the proportion of ATP turnover in total
O2 of hepatopancreas cells
was slightly lower (44–45%) due to a higher percentage of
non-mitochondrial respiration.
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Mitochondrial abundance and fractional volume
Mitochondrial abundance was similar in gills and hepatopancreas
(F1,354=1.54, P=0.2148), whereas mitochondrial
fractional volume was nearly twice as high in hepatopancreas than in gills
(F1,356=34.15, P<0.0001,
N=29–31), indicating the larger size of hepatopancreas
mitochondria (Tables 3,
4). Cd exposure resulted in a
decrease in mitochondrial numbers in both organs that was statistically
significant in gills and marginally significant in hepatopancreas
(F1,172=4.50, P=0.03, N=29–31 and
F1,174=2.68, P=0.10, N=29–31,
respectively). This decrease was more pronounced at higher temperatures so
that mitochondrial numbers dropped by 8–13% in Cd-exposed oysters at
12°C and by 20–24% at 28°C
(Table 3). Mitochondrial volume
was significantly reduced in Cd-exposed oysters in hepatopancreas
(F1,177=4.60, P=0.03, N=29–31). In
gills, interaction between temperature and Cd exposure had a significant
effect on mitochondrial fractional volume (P=0.015), thus preventing
analysis of the effects of single factors. Therefore, effects of Cd exposure
on the mitochondrial volume in gills were analyzed separately for each
acclimation temperature; Cd effects were significant at 20°C
(P=0.008) but not at 12°C or 28°C (P=0.58 and 0.24,
respectively) (Table 4).
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O2 in oysters at 12 and
20°C, indicating elevated energy demand. By contrast, at 28°C there
was no increase in cellular
O2 in response to Cd. This
agrees with our earlier studies showing elevated whole-organism SMR in
Cd-exposed oysters at 20°C but not at 28°C
(Lannig et al., 2006) and
suggests good correspondence between cellular and whole-organism metabolic
response to Cd. Surprisingly, mitochondrial oxidation capacity (measured at
the respective acclimation temperature) did not increase with increasing
temperature of acclimation and was highest in 20°C-acclimated oysters
compared with the 12°C- or 28°C-acclimated ones. In fact,
mitochondrial capacity for ATP synthesis was >2 times lower in
28°C-acclimated oysters than in their 20°C-acclimated counterparts
despite the fact that the whole-cell oxygen demand increased nearly twofold at
28°C. This indicates that oyster mitochondria may function close to their
capacity limits at 28°C and could explain why SMR does not further
increase in response to Cd at this temperature.
Analysis of cellular energy budgets in oysters showed that mitochondrial
proton leak and protein synthesis represent major energy costs, accounting for
22–38% and 6–17% of the total oxygen consumption, respectively.
These estimates are similar to the values reported in other poikilotherms (for
reviews, see Brand et al.,
1991; Hand and Hardewig,
1996; Hulbert and Else,
1999; Hulbert et al.,
2002). An increase in acclimation temperature did not
significantly affect fractions of the total energy spent on proton leak,
protein synthesis, overall ATP turnover or non-mitochondrial respiration
despite an overall increase in cellular energy demand. This indicates a
concerted change in the rates of different energy-requiring cellular processes
during temperature acclimation in oysters. By contrast, Cd exposure had a
disproportionate effect on some components of the energy budget (particularly
protein synthesis and, to a lesser extent, proton leak) while not affecting
the others (non-mitochondrial respiration and overall ATP turnover). Cd
exposure resulted in a strong, 1.5-fold increase in mitochondrial proton leak
in gill and hepatopancreas cells in 12°C-acclimated oysters, whereas no
change was observed at 20 and 28°C. This pattern closely corresponds to
the data on isolated mitochondria, where Cd exposure resulted in a 1.5-fold
increase in proton leak at 12°C but not 20 or 28°C. Thus, elevated
proton leak may contribute to the observed increase in energy demand of
Cd-exposed gill cells at 12°C.
Our data show that an increase in the cost of protein synthesis was the
most consistent metabolic response to Cd, indicating that this mechanism may
significantly contribute to the Cd-induced increase in energy demand. In
gills, prolonged Cd exposure resulted in a >6-fold and >2-fold increase
in the protein synthesis costs at 12 and 20°C, respectively. In Cd-exposed
hepatopancreas cells, energy demand for protein synthesis increased by a
factor of 2–2.5 across all studied temperatures. This increase is likely
to reflect elevated rates of protein synthesis, which may include protein
deposition (growth) as well as proteins synthesized for Cd detoxification,
general stress protection and replacement of the damaged housekeeping
proteins. There is no evidence that Cd exposure can result in elevated rates
of protein deposition and growth; in fact, Cd exposure is known to stunt
growth and lead to protein loss in oysters, especially at high Cd levels
(reviewed in Roesijadi, 1996).
Notably, this study also showed that cellular protein content decreased by
approximately 10% in gills and did not change in hepatopancreas in Cd-exposed
oysters, indicating that there was no increase in protein deposition in
response to Cd in oyster cells.
Overexpression of inducible stress proteins is a particularly attractive
hypothesis to explain the elevated rates of protein synthesis in response to
Cd exposure. It has been shown that Cd exposure results in elevated expression
of metallothioneins (MTs) and heat-shock proteins (HSPs) in oysters
(Roesijadi et al., 1997;
Piano et al., 2004;
Moraga et al., 2005). Due to
the short half-life of MTs and HSPs
(Andersen et al., 1978;
Mehra and Bremner, 1985;
Pelham, 1990), high synthesis
rates would be required to maintain elevated steady-state levels of these
proteins in Cd-exposed oysters. On the other hand, Cd exposure can induce
oxidative damage of proteins (Sandalio et
al., 2001; Collen et al.,
2003; Shi et al.,
2005) and/or disrupt their native structure due to binding to the
critical thiol and imidazol groups (reviewed in
Valko et al., 2005). Thus,
synthesis costs to replace the damaged housekeeping proteins may also add to
the elevated energy demand in Cd-exposed oysters. Quantification of the
relative contributions of stress protein expression and the replacement of the
damaged housekeeping to the global costs of protein synthesis is an important
question that requires further investigation in order to further our
understanding of the mechanisms of metabolic response to stress.
It is worth noting that the above interpretation (i.e. that elevated
protein synthesis costs in Cd-exposed oysters are reflective of the elevated
rates of protein synthesis) assumes that the efficiency of protein synthesis
(i.e. ATP demand per peptide bond) is the same in control and Cd-exposed
oysters. If this does not hold true and the efficiency of the protein
synthesis is adversely affected by Cd, an increase in energy demand for
protein synthesis may reflect higher ATP demand per unit synthesized protein
rather than elevated rates of synthesis. Few studies that have directly
measured protein synthesis efficiency in marine invertebrates indicate that
ATP demand per peptide bond is fairly constant over a broad range of
environmental temperatures and at different stages of life cycles
(Storch and Pörtner,
2003; Pace and Manahan,
2006). Unfortunately, the effects of Cd exposure on the ATP demand
for protein synthesis have not yet been studied and clearly warrant further
investigation. Irrespective of the molecular mechanisms, elevated energy
expenditure for protein synthesis could divert energy from other
energy-demanding cellular processes; indeed, the fraction of protein synthesis
costs in the total ATP turnover increased by approximately 20% in Cd-exposed
oysters.
The potential impact of the stress-induced energy demand would depend on
the ability of the metabolic machinery to adequately increase energy output to
compensate for it. Obviously, if the energy supply is adequate, elevated
energy demand per se may not cause any adverse effects at the
whole-organism level. Our data suggest that this is not the case in oysters
exposed to combined Cd and temperature stress. No compensatory increase in
mitochondrial oxidation rate, abundance or fractional volume was observed in
Cd- and/or temperature-stressed oysters, indicating that the supply side of
the cellular energy budget may become limiting under combined exposure to
those stressors. In fact, there was a trend for decreased mitochondrial
abundance in Cd-exposed oysters compared with their control counterparts, and
this decrease was most pronounced in warm-acclimated oysters that experience
the highest energy demand. Our earlier studies also showed that oyster
mitochondria become increasingly more sensitive to the toxic effects of
cadmium as the environmental temperature rises
(Sokolova, 2004). Taken
together, these data indicate that combined exposure to temperature and Cd
stress may result in a decreased mitochondrial capacity of oyster tissues,
which, in conjunction with elevated SMR, may narrow the aerobic scope in
oysters.
As a corollary, our data show that Cd exposure and elevated temperatures
strongly affect both the demand side and the supply side of cellular energy
budgets in the model marine poikilotherm Crassostrea virginica.
Elevated cellular energy demands in Cd-exposed oysters are due to the
increased costs of protein turnover and, at cold acclimation temperatures, are
also partially due to the elevated mitochondrial proton leak. Cellular aerobic
capacity does not increase in parallel to compensate for elevated energy
demand. This may result in a discrepancy between energy demand and energy
supply and lead to the reduced aerobic scope for activity, growth and/or
reproduction and thus to potential fitness costs in temperature- and
Cd-stressed oysters. The disconnection between indices of energy supply and
demand also has important methodological implications, suggesting that
demand-side and supply-side endpoints should be considered separately in
studies of energy budgets. It strongly cautions against the use of capacity
indices such as the maximum activity of electron transfer system as a measure
of energy demand, as was proposed recently
(Fanslow et al., 2001;
De Coen and Janssen, 2003;
Smolders et al., 2004). Our
study shows that both elevated temperatures and Cd exposure lead to high
energy demand and adversely affect mitochondrial aerobic capacity despite
different molecular mechanisms of action. This could explain strong synergism
between these environmental stressors and suggests that elevated temperatures
(as expected during seasonal acclimatization and/or global climate change) may
strongly enhance pollution-related stress in marine poikilotherms.
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Acknowledgments |
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